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Antimicrobial activity of menthol modified nanodiamond particles
Volodymyr Turcheniuk, Viktoria Raks, Rahaf Issa, Ian R. Cooper, Peter J. Cragg , Roxana Jijie, Nicoleta
Dumitrascu, Lyuba I. Mikhalovska, Alexandre Barras, Vladimir Zaitsev, Rabah Boukherroub, Sabine
Szunerits
Abstract
Advances in nanotechnology have seen the development of several microbiocidal nanoparticles
displaying activity against biofilms. These applications benefit from one or more combinations of the
nanoparticle properties. Nanoparticles may indeed concentrate drugs on their surface resulting in
polyvalent effects and improved efficacy to fight against bacteria. Nanodiamonds (NDs) are among
the most promising new materials for biomedical applications. We elucidate in this paper the effect
of menthol modified nanodiamond (ND-menthol) particles on bacterial viability against Grampositive (Staphylococcus aureus) and Gram-negative (Escherichia coli) bacteria. We show that while
ND-menthol particles are non-toxic to both pathogens, they show significant antibiofilm activity. The
presence of ND-menthol particles reduces biofilm formation more efficiently than free menthol,
unmodified oxidized NDs and ampicillin, a commonly used antibiotic. Our findings might be thus a
step forward towards the development of alternative non antibiotic based strategies targeting
bacterial infections.
Keywords
Nanodiamond particles; Menthol; Functionalization; Antimicrobial activity; Biofilm inhibition
1. Introduction
Bacterial infections are one of the most serious risks to public health. Extensive efforts have been
devoted to develop rapid and sensitive methods for the detection, identification of the pathogen
strain at hand and for destruction or inhibition of pathogens and their actions [1], [2], [3] and [4].
Despite considerable recent progress in the understanding of the mechanisms underlying bacterial
infections, and in the development of nanostructured materials displaying antibacterial properties
and activity against biofilms [1], [5], [6], [7] and [8], the quest to design and fabricate new
antibacterial nanostructures remains a high research priority. Nanoparticles have been considered
as affective solution to fight against bacterial infections [4], [9], [10], [11], [12], [13], [14] and [15].
Such nanostructures allow the concentration of antibacterial agents and functions on their surface
to deliver polyvalent effects. Some materials such as silver are themselves antibacterial and can
result in enhanced efficacy [16], [17] and [18]. Although recent studies using silver nanoparticles
appear promising [19], the antimicrobial activity is highly dependent upon their size and shape, with
smaller particles demonstrating greater activity [20] and [21]. More recently concerns about the
cytotoxic effects of silver nanoparticles against human cells have been voiced [22]. Moreover,
practical applications of Ag NPs are often hampered by the aggregation and loss of antibacterial
activity [23].
Among the potential alternatives to these compounds are nanodiamond particles (NDs). One of the
advantages of NDs over other carbon-based materials such as fullerenes and carbon nanotubes is
that they are completely inert, optically transparent, biocompatible and can be functionalized in
many ways depending on their intended ultimate application [24], [25], [26], [27], [28], [29], [30] and
[31]. Although their in vivo toxicity depends on their particular surface characteristics [32], ND
particles do not induce significant cytotoxicity in a variety of cell types [32], [33], [34] and [35] and
have been used in a variety of biomedical applications. More recently, the antibacterial activity of
diamond particles [36], [37] and [38] and their potential to interfere with biofilm formation [1] have
been highlighted. The underlying anti-adhesive strategy proposed was based on the interfering with
type 1 fimbriae-mediated mannose recognition events [39]. Such biofilm disrupting activity had not
been observed previously for other glyco-nanoparticles (glyco-NPs) such as glycofullerenes, goldbased glyco-NPs or for other multivalent mannose-derived molecules [2] and [40]. Wehling et al.
have concluded that the bactericidal activity of diamond particles is directly linked to their surface
chemistry, being thus the driving force of antibacterial effects [37].
In this paper, we investigate the antimicrobial properties of ND particles modified with menthol (Fig.
1A) and their potential to inhibit the growth of Gram-positive (Staphylococcus aureus) and Gramnegative (Escherichia coli) bacteria.
Fig. 1. (A) Schematic illustration of the stepwise chemical functionalization of nanodiamond particles
(ND) to give the target ND-menthol cluster; (B) synthetic route to menthol derivative (2): (i) succinic
anhydride, 4-DMAP, CHCl3, reflux, overnight, 84%; (ii) NHS, DCC, CH2Cl2, r.t, overnight, 95%.
Menthol, also known as mint camphor, is a naturally occurring cyclic terpene alcohol of plant origin,
which has been used since antiquity for medicinal purposes [41], [42] and [43]. It is a major
constituent in the essential oils of Mentha canadensis L. (cornmint) and peppermint possessing wellknown cooling characteristics and a residual minty smell of the oil remnants from which it was
obtained. Several studies have additionally demonstrated that next to the effect of sensory
parameters, menthol has antibacterial and antifungal activities [44] and [45]. However, the
antimicrobial activity was reported to be lower compared to antibiotics such as penicillin [46]. More
recently, Imbert and co-workers showed the in vitro activity of terpenes, including that of menthol,
against Candida biofilms [47]. We show here that ND-menthol particles affect moderately S. aureus
and E. coli growth in a concentration dependent manner. In addition, the ND-menthol particles are
found to inhibit S. aureus and E. coli-driven biofilm growth significantly.
2. Experimental
2.1. Materials
Hydroxyl-terminated nanodiamond (ND-OH) particles were purchased from International
Technology Center (Raleigh, NC, USA) and exhibit a primary average particle size of 4.0 nm.
(1R,2S,5R)-(−)-menthol, 4-dimethylaminopyridine (4-DMAP), succinic anhydride, hexane (Hex),
dichloromethane (CH2Cl2), chloroform (CHCl3), ethyl acetate (EtOAc), N,N′-dicyclohexylcarbodiimide
(DCC) and N-hydroxysuccinimide (NHS) were purchased from Sigma-Aldrich and used as received. All
microbiological media, equipment and the ampicillin were purchased from Fisher Scientific (UK).
2.2. Synthesis of menthol derivative (2)
2.2.1. Succinic acid mono-(2-isopropyl-4-methyl-cyclohexyl) ester (1)
To the solution of menthol (0.8 g, 5 mmol) in chloroform, 4-DMAP (0.31 g, 2.5 mmol) and succinic
anhydride (1.02 g, 10 mmol) were added. This mixture was stirred overnight under reflux. After
solvent evaporation, the crude product was purified using flash column chromatography with
Hex:EtOAc as eluent to give white crystals 1.1 g (84%). 1H NMR (300 MHz, CDCl3) δ 12.2–11.1
(broad, 1H) 4.77–4.68 (m, 1H), 2.67–2.55 (m, 4H), 2.05–1.95 (m, 1H), 1.91–1.79 (m, 1H), 1.74–1.62
(m, 2H), 1.55–1.33 (m, 2H), 1.1–0.8 (m, 10H), 0.78–0.7 (d, 3H).
2.2.2. Succinic acid 2,5-dioxo-pyrrolidin-1-yl ester 2-isopropyl-4-methyl-cyclohexyl ester (2)
To the solution of acid (1) (0.5 g, 1.9 mmol) in dichloromethane, N,N′-dicyclohexylcarbodiimide (0.4
g, 1.9 mmol) and N-hydroxysuccinimide (0.22 g, 1.9 mmol) were added and the mixture was stirred
overnight at room temperature. After filtration and solvent evaporation a white powder was
obtained, yield: 95%. 1H NMR (300 MHz, CDCl3) δ 4.77–4.68 (m, 1H), 2.85–2.81 (s, 4H), 2.54–2.46
(m, 4H), 2.01–1.94 (m, 1H), 1.88–1.76 (m, 1H), 1.73–1.62 (m, 2H), 1.55–1.33 (m, 2H), 1.22–0.82 (m,
10H), 0.75–0.7 (d, 3H).
2.3. Preparation of menthol-modified NDs (ND-menthol)
Menthol derivative (2) (10 mM), was dissolved in 5 mL water and added to a suspension of ND-OH
particles in water (10 mg in 5 mL). The solution was stirred at room temperature for 24 h under
nitrogen. The resulting ND-menthol particles were isolated by centrifugation at 10,000 rpm, purified
through four consecutive wash/centrifugation cycles at 10,000 rpm with ethanol, and water and
finally oven dried at 50 °C for 24 h.
2.4. Characterization
2.4.1. FTIR spectroscopy
Fourier transformed infrared (FTIR) spectra in transmission mode were recorded using a
ThermoScientific FTIR instrument (Nicolet 8700) with a resolution of 4 cm− 1. Dried ND powder (1
mg) was mixed with KBr powder (100 mg) in an agate mortar. The mixture was pressed into a pellet
under 10 ton load for 2–4 min, and the spectrum was recorded immediately. Sixteen accumulative
scans were collected. The signal from a pure KBr pellet was subtracted as a background.
2.4.2. Particle size measurements
ND suspensions (20 μg mL− 1) in water were sonicated. The particle size of the ND suspensions was
measured at 25 °C using a Zetasizer Nano ZS (Malvern Instruments S.A., Worcestershire, U.K.) in 173°
scattering geometry and the zeta potential was measured using the electrophoretic mode.
2.4.3. Transmission electron microscope (TEM)
TEM measurements were performed in a FEI Tecnai G2 20 equipped with EDS micro-analysis, Gatan
energy filter (EELS), electron precision and tomography.
2.4.4. Thermogravimetric analysis (TGA)
Thermogravimetric analysis (TGA) measurements were made in Al2O3 crucibles in an atmosphere of
nitrogen at a heating rate of 10 °C min− 1 using a TA Instruments Q50 thermogravimetric analyzer.
2.5. Antimicrobial assays
2.5.1. Bacterial biofilm formation
Overnight cultures of E. coli (NCTC 8196) and S. aureus (NCTC 6571) were prepared by inoculating 10
mL of nutrient broth and incubating at 37 °C for 18 h on a rotary shaker (Stuart Scientific) at 120
rpm. After incubation, the cultures were diluted to OD600 of 0.1 in nutrient broth at a final volume
of 100 mL, and a 96-well plate was set up as follows. Each assay sample required 6 wells, with
another 6 wells for a negative control/blank, which consisted of 125 μL of sterile nutrient broth.
Next, 110 μL of culture plus 15 μL of PBS was pipetted into the next 6 wells to act as the positive
growth control. In the remaining wells, the assay was set up as follows: 110 μL of culture and 15 μL
of the appropriate NDs and ampicillin solutions. Sterile nutrient broth and NDs and sterile nutrient
broth and ampicillin were also set up in sets of six wells each to act as controls. The 96-well plate
was incubated at 37 °C for 24 h with no agitation. After 24 h, the plate was removed from the
incubator, and the supernatant of each well was removed by pipetting. The plates were
subsequently allowed to dry for 15–20 min in a Class Two Microbiological Hood. Next, 125 μL of PBS
was transferred into each well in order to remove any non-adhering cells. The PBS was then
removed by pipetting, and the plates were again allowed to dry in the Class Two Microbiological
Hood for 15 min. All wells were stained by adding 150 μL of 0.9% crystal violet in sterile reverse
osmosis water, and the plates were incubated at room temperature for 15 min. After this time, the
stain was removed by pipetting, and excess dye was washed off by manually adding and then
removing 165 μL of PBS. To solubilize the crystal violet, 175 μL of 95% ethanol was added to each
well, and the plates were incubated at room temperature for 30 min. The bacterial cell density was
determined by the optical density of each well at 595 nm, using the negative control wells as blanks.
2.5.2. Viability of bacterial biofilms
The LIVE/DEAD® BacLight™ assay kit was used to determine the effect of ND-menthol (100 μg/mL)
on bacterial cell viability. Overnight cultures of E. coli and S. aureus were prepared by inoculating 10
mL of nutrient broth and incubating at 37 °C for 18 h on a rotary shaker (Stuart Scientific) at 120
rpm, and subsequently diluted to OD600 of 0.01 in nutrient broth. ND-menthol (100 μg/mL) was
suspended in sterile nutrient broth and added to respective cultures. The cultures were incubated in
the presence of ND-methanol for 3 h (logarithmic growth phase) at 37 °C at 120 rpm. After
incubation, the cultures were centrifuged at 10,000 rpm for 5 min, and the supernatant solution was
discarded. The resulting cell pellets were washed with 1.5 mL sterile PBS, and centrifuged at 10,000
rpm for 5 min, three times to remove media constituents that may interfere with the stain. After
each wash the supernatant was discarded and the pellet collected. After the final wash in PBS, the
cells were re-suspended in 1 mL PBS. On ice, equal volumes of component A and component B of the
LIVE/DEAD® BacLight™ stains were added to the cell suspensions and thoroughly mixed by vortexing.
Next, 3 μL of stain was added to 1 mL of culture suspension and the samples were incubated in the
dark at room temperature for 20 min. Finally, 5 μL of the stained bacterial suspension was captured
between a clean microscope slide and an 18 mm coverslip. Samples were visualized using a Leica TCS
SP5 Confocal Laser Scanning Microscope.
3. Results and discussion
Covalent linking of antibacterial agents to surfaces has been considered as a reliable strategy to
design antibacterial materials [48] and [49]. We opted here for the covalent linking of a carboxylic
acid modified menthol derivative (1) to the hydroxyl groups of ND particles through an esterification
reaction with the aim to determine the antibacterial properties of the menthol modified ND particles
(Fig. 1). The successful integration of the menthol moiety onto ND was evidenced using Fourier
transform infrared spectroscopy (FTIR) (Fig. 2A). ND-OH particles show a broad peak at 3400 cm− 1
assigned to the vibration of surface hydroxyl groups or/and adsorbed water molecules, and an
additional sharper one at 1633 cm− 1 due to the bending mode δ(OH) of surface hydroxyl groups on
the NDs. In addition, the band at 1107 cm− 1 is indicative of the presence of C–O–C– functions of
cyclic ethers. The FTIR spectrum of the ND-menthol particles displays in addition to bands at 3400
cm− 1 (vibration of surface hydroxyl groups or/and adsorbed water molecules) 1633 cm− 1 (bending
mode of surface hydroxyl groups) and 1107 cm− 1 is (C–O–C– functions of cyclic ethers), a band at
1717 characteristic of the Cdouble bond; length as m-dashO stretching of the installed ester
function. The C–H stretching vibration modes in menthol are seen between 2850 and 2970 cm− 1.
Fig. 2. (A) FTIR spectra of ND-OH (black) and ND-menthol (red); (B) transmission electron microscopy
(TEM) images of ND-menthol particles together with histogram of particle size distribution; (C)
thermogravimetric analysis of menthol derivative (2) (green), ND-menthol (red) and ND-OH (black).
TEM images of ND-menthol particles (Fig. 2B) reveal the presence of spherical particles with a size
distribution in the range of 15 ± 5 nm in diameter. The hydrodynamic diameter of ND-menthol
particles is 92 ± 10 nm, comparable to that of ND-OH (89 nm) with a zeta potential of 29.5 ± 0.5 mV,
slightly lower than that of ND-OH (35.3 ± 1.6 mV) (Table 1).
Table 1. Physico-chemical properties of ND-OH and ND-menthol.
Hydrodynamic diameter
(nm)
PI
ND-OH
89 ± 13
0.246 ± 0.002 35.3 ± 1.6
NDmenthol
92 ± 10
ND structure
Zeta potential
(mV)
0.232 ± 0.015 29.5 ± 0.5
To gain more insight onto the strength of the bonding between menthol derivative (2) to the surface
of ND-OH particles, thermogravimetric analysis of ligand (2), ND-OH and ND-menthol particles was
performed (Fig. 2C). The TGA thermogram of the as-received ND-OH particles indicates that they are
stable up to 550 °C without any apparent mass loss [50] and [51]. TGA measurements indicate a
weight loss of ≈ 7.45% above 95 °C mainly due to absorbed water [50]. Menthol derivative (2)
decomposes beyond ≈ 150 °C. The removal of the covalently linked menthol derivative (2) shows its
decomposition at temperatures above ≈ 200 °C. The weight loss between 200 and 300 °C
corresponding to the removal of the menthol acid (1) is around 58% indicating a surface loading of
menthol derivative (2) of 2.1 ± 0.2 mmol g− 1.
To test the effect of ND-menthol particles on the growth of pathogens and bacteria induced biofilm
formation, Gram-positive S. aureus and Gram-negative E. coli were used as model organisms. S.
aureus are extremely versatile pathogenic bacteria that cause a wide range of syndromes, ranging
from minor skin and soft tissue infections to life-threatening pneumonia [52] and [53]. Gramnegative E. coli bacteria, widely used as a control organism in microbiological assays, are present
within the gastro-intestinal (GI) tract, or as an intra- or extra-intestinal pathogen. A number of
antibiotics in use today are delivered orally, and hence the ability of a compound to effect the
normal GI flora is of prime importance. Disruption of the GI flora is not uncommon, but the complete
removal of strains or a significant alteration in the relative proportions of the constituent organisms
might lead to opportunistic infection by E. coli, or other organisms. Key virulence factors associated
with this organism are attachment pili which facilitate adherence to the mucosal lining of the GI or
urinary tract [54].
To determine the extent of the antimicrobial activity of ND-menthol particles, they must be assessed
against those of known cytotoxic agents to give a positive control. An antibiotic sensitivity test was
conducted using a range of 10 common antibiotics at a concentration of 1 μg mL− 1. The behavior of
the different antibiotics towards E. coli growth was determined by the standard microbiological
technique of measuring the size of the zone of inhibition around the point where the antibiotic was
placed ( Fig. 3). Extremely weak inhibition was observed for penicillin G, while ciprofloxacin and
ceftiofur showed relatively strong activity. From the results of the sensitivity test, ampicillin was
chosen as a classical antibiotic with an intermediate effect and used for comparison in further
experiments.
Fig. 3. Antibiotic action on E. coli growth on agar plates.
The planktonic growths of S. aureus and E. coli were thus followed over a 6 hour period in the
absence and presence of the ND-menthol particles at different concentrations (Fig. 4). As control the
effect of bacterial growth in the presence of unmodified ND-OH (10 μg mL− 1) and ampicillin (10 μg
mL− 1) was investigated. As shown in Fig. 4A, while ND-OH particles do not influence the planktonic
growth of S. aureus, ND-menthol particles had a concentration dependent effect on the growth of S.
aureus by moderately reducing the growth of the Gram-positive pathogen. However, even at the
highest concentration test, the growth of S. aureus was significantly less effective than when an
antibiotic was present. In the case of E. coli a similar behavior was observed ( Fig. 4B). While for the
first 2 h, E. coli growth was inhibited in the presence of even low concentrations of ND-menthol,
their efficiency started to be compromised partially thereafter allowing the planktonic growth of E.
coli. The effect of ND-menthol on the growth of S. aureus and E. coli was evaluated using a
LIVE/DEAD test (Invitrogen) following the manufacturer's instructions. In Fig. 5 show cell growth for
S. aureus and E. coli in the absence (control) and presence of ND-menthol (100 μg/mL) and after
being stained with the LIVE (green)/DEAD (red) stains. These data suggest that ND-menthol particles
do not alter the bacterial viability of S. aureus and E. coli. and are thus non toxic. The toxic effects on
membrane structures and function have generally been used to explain the antimicrobial activity of
menthol, although the exact mechanism of action is not fully understood [44]. Menthol is believed
to alter cell permeability by penetrating between the fatty acid chains making up the membrane
lipid bilayers, disrupting lipid packaging and changing membrane fluidity. These phenomena result in
major surface alterations and morphological modifications of Gram-positive and Gram-negative
bacteria and in leakage of intracellular materials. Linking of menthol to ND particles seems to limit
the bactericidal activity of menthol. It is indeed in general believed, that the antibacterial effect of
diamond particles is related to its zeta-potential, where particles with positive potential
demonstrated no effect, but those with negative zeta potential were shown to disrupt the cell wall
and cytoplasmic membrane to release the cytoplasm from the cell [55]. ND-menthol as well as NDOH particles used as control have both positive zeta-potential (Table 1). More recently, Wehling et
al. [37] studied the bactericidal activity of partially oxidized NDs. Their experiments suggested that
the antibacterial activity of NDs is linked to the presence of partially oxidized and negatively charged
surfaces, specifically those containing acid anhydride groups. From the FTIR image of ND-OH and
ND-menthol particles (Fig. 2A), the presence of acid anhydride groups with an absorption band at
around 1850–1750 cm− 1 is not visible explaining in addition to the presence of viable bacteria.
Fig. 4. Growth curves of (A) S. aureus and (B) E. coli in the presence of ND-OH (10 μg mL− 1),
ampicillin (1 μg mL− 1) and ND-menthol at 1, 10 and 100 μg mL− 1.
Fig. 5. Viability of S. aureus and E. coli imaged by the LIVE/DEAD® BacLight™ assay (× 40
magnification) (a) control (no NDs present); (b) ND-menthol. Green fluorescence indicates live cells
while red or yellow fluorescence indicates dead or compromised cells.
While ND-menthol particles were found to not interfer with the development of the planktonic
growth of S. aureus and E. coli, we had reasons to believe that ND-menthol particles might affect the
adhesion capacitance of the pathogens [47]. The potential of the different nanostructures to inhibit
S. aureus and E. coli biofilm formation was furthermore investigated. In the case of S. aureus, NDmenthol particles exhibited dose-dependent inhibition of biofilm formation as shown in Fig. 6A. In all
cases strong biofilm inhibition is observed with ND-menthol particles. At a particle concentration of
100 μg mL− 1 the inhibition of biofilm formation was comparable to that of the antibiotic, ampicillin,
used. Free menthol (100 μg mL− 1), on the other hand, did not show any significant biofilm
inhibition. ND-OH particles exhibited a moderate biofilm inhibition even at higher concentrations.
The reason for the increased biofilm inhibition effect of ND-menthol over free menthol is most likely
put down to their more efficient harnessing of a multivalent effect. The difference is probably also
driven, in part, by the intrinsic physiochemical properties of the NDs. While more detailed studies
are needed to understand fully the antibiofilm activity of menthol modified NDs, the reported
change in bacteria membrane fluidity in the presence of menthol [44] seems to reduce the
adherence capacity of S. aureus. This is also valid for E. coli biofilms. In the case of E. coli, the NDmenthol particles rival with that of ampicillin even at the lowest concentration level ( Fig. 6B), with
high ND-menthol concentrations resulted in strong biofilm inhibition.
4. Conclusion
In this work, we have demonstrated that menthol-conjugated nanodiamond particles have marked
effects on S. aureus and E. coli-mediated biofilm formation. The conjugation strategy developed
based on the formation of an ester bond between the hydroxyl groups of ND and the carboxylic acid
group of a synthetic menthol derivative (1) proved to be highly efficient for the coupling of the
antibacterial agent. The demonstration that ND-menthol particles are able to effectively impede
bacterial adhesion of Gram-positive and Gram-negative bacteria is consistent with our earlier
findings that integration of a biofilm inhibitor onto diamond particles has marked E. coli antiadhesive activity. In addition, neither ND-OH nor menthol controls are seen to show any antiadhesive activity, underlying that the observed activities are specific to ND-menthol. Moreover, NDmenthol showed better biofilm inhibition than ampicillin even at low particle concentrations. Taken
together, the data support that the presence of menthol moiety is the primary ingredient driving the
biofilm-inhibitory activity observed for the ND-menthol conjugate.
Fig. 6. Biofilm formation of (A) S. aureus and (B) E. coli in the absence and presence of ND-menthol
(1, 10, 100 μg mL− 1), ND-OH (1, 10, 100 μg mL− 1), ampicillin (1 μg mL− 1) and menthol (100 μg mL−
1).
Prime novelty statement
1. Diamond nanoparticles modified with menthol induce reduction of biofilm formation more
efficiently than free menthol.
2. ND-menthol particles also compete with ampicillin, a commonly used antibiotic.
3. Our findings might be thus a step forward towards the development of alternative non-antibiotic
based strategies targeting bacterial infections.
Acknowledgments
V.T., R.B. and S.S. gratefully acknowledge the financial support from the Centre National de
Recherche Scientifique (CNRS), the Université Lille 1, the Nord Pas de Calais region and the Institut
Universitaire de France (IUF). R.I. thanks the University of Brighton for a studentship. V.R. and V.C.
acknowledge the financial support of the European Commission (IRSES project: 269009). We also
acknowledge the support from the European Union through the FP7-PEOPLE-2010-IRSES action
“Photorelease” (grant number 269099). R. J. thanks Institut Français in Romania for a PhD
scholarship. We thank Profs P. Woisel, J. Lyskawa and A. Malfait (UMET UMR CNRS 8207, Université
de Lille Nord de France, ENSCL) for the technical support with TGA measurements.
References
[1] A. Barras, F.A. Martin, O. Bande, J.S. Baumann, J.-M. Ghigo, R. Boukherroub, C. Beloin, A.
Siriwardena, S. Szunerits, Glycan-functionalized diamond nanoparticles as potent E. coli antiadhesives, Nanoscale, 5 (2013), p. 2307
[2] M. Durka, K. Buffet, J. Iehl, M. Holler, J.F. Nierengarten, J. Taganna, J. Bouckaert, S.P. Vincent, The
functional valency of dodecamannosylated fullerenes with Escherichia coli FimH—towards novel
bacterial antiadhesives, Chem. Commun., 47 (2011), p. 1321
[3] P. Laurino, R. Kikkeri, N. Azzouz, P.H. Seeberger, Detection of bacteria using glyco-dendronized
polylysine prepared by continuous flow photofunctionalization, Nano Lett., 11 (2011), p. 73
[4] P.C. Ray, S.A. Khan, A.K. Singh, D. Senapati, Z. Fan, Nanomaterials for targeted detection and
photothermal killing of bacteria, Chem. Soc. Rev., 41 (2012), p. 3193
[5] A. Herman, A.P. Herman, Nanoparticles as antimicrobial agents: their toxicity and mechanisms of
action, J. Nanosci. Nanotechnol., 14 (2014), pp. 946–957
[6] R.P. Allaker, K. Memarzadeh, Nanoparticles and the control of oral infections, Int. J. Antimicrob.
Agents, 43 (2014), pp. 95–104
[7] S. Chernousova, M. Epple, Silver as antibacterial agent: ion, nanoparticle, and metal, Angew.
Chem. Int. Ed., 52 (2012), pp. 1636–1653
[8] M.R. Das, R.K. Sarma, R. Saikia, V.S. Kale, M.V. Shelke, P. Sengupta, Synthesis of silver
nanoparticles in an aqueous suspension of graphene oxide sheets and its antimicrobial activity,
Colloids Surf. B Biointerfaces, 83 (2011), p. 16
[9] Y. Zhao, Y. Tian, Y. Cui, L. Wenwen, W. Ma, X. Jiang, Small molecule-capped gold nanoparticles as
potent antibacterial agents that target gram-negative bacteria, J. Am. Chem. Soc., 132 (2010), p.
12349
[10] O. Akhavan, R. Azimirad, S. Safed, E. Hassani, CuO/Cu(OH)2 hierarchical nanostructures as
bactericidal photocatalysts, J. Mater. Chem., 21 (2011), p. 9634
[11] G. Qi, L. Li, F. Yu, H. Wang, Vancomycin-modified mesoporous silica nanoparticles for selective
recognition and killing of pathogenic gram-positive bacteria over macrophage-like cells,ACS Appl.
Mater. Interfaces, 5 (2013), p. 10874
[12] R.Y. Pelgrift, A.J. Friedman, Nanotechnology as a therapeutic tool to combat microbial
resistance, Adv. Drug Deliv. Rev., 65 (2013), pp. 1803–1815
[13] K. Forier, K. Raemdonck, S.D. De Smedt, J. Demeester, T. Coenye, K. Braeckmans, Lipid and
polymer nanoparticles for drug delivery to bacterial biofilms, J. Control. Release, 190 (2014), pp.
607–623
[14] X. Zhu, A.F. Radovic-Moreno, J. Wu, R. Langer, J. Shi, Nanomedicine in the management of
microbial infection — overview and perspectives, Nano Today, 9 (2014), pp. 478–498
[15] P. Ma, R.J. Mumper, Anthracycline nano-delivery systems to overcome multiple drug resistance:
a comprehensive review,Nano Today, 8 (2013), p. 313
[16] M.R. Das, R.K. Sarma, S.C. Borah, R. Kumri, R. Saikia, A.B. Deshmukh, M.V. Shelke, P. Sengupta,
S. Szunerits, R. Boukherroub, Synthesis of silver nanoparticles in an aqueous suspension of graphene
oxide sheets and its antimicrobial activity, Colloids Surf. B, 105 (2013), pp. 128–136
[17] J.S. Kim, E. Kuk, K.N. Yu, J.H. Kim, S.J. Park, H.J. Lee, S.H. kim, Y.K. Park, Y.H. Park, C.Y. Hwang,
Y.K. Kim, Y.S. Lee, D.H. Jeong, M.H. Cho, Antimicrobial effects of silver nanoparticles, Nanomedicine,
3 (2007), pp. 95–101
[18] S. Mohanty, S. Mishra, P. Jena, B. Jacob, B. Sarkar, A. Sonawane, An investigation on the
antibacterial, cytotoxic, and antibiofilm efficacy of starch-stabilized silver nanoparticles,
Nanomedicine, 8 (2012), pp. 916–924
[19] J.-H. Liu, D.A. Sonshine, S. Shervani, R.H. Hurt, Controlled release of biologically active silver
from nanosilver surfaces, ACS Nano, 4 (2010), p. 6903
[20] A. Panacek, L. Kvitek, R. Prucek, M. Kolar, R. Vecerova, N. Pizurova, V.K. Sharma, T. Nevecna, R.
Zboril, Silver colloid nanoparticles: synthesis, characterization, and their antibacterial activity, J.
Phys. Chem. B, 110 (2006), p. 16248
[21] S. Pal, Y.K. Tak, J.M. Song, Does the antibacterial activity of silver nanoparticles depend on the
shape of the nanoparticle? A study of the gram-negative bacterium Escherichia coli, Appl. Environ.
Microbiol., 73 (2007), pp. 1712–1720
[22] K.K.Y. Wong, X. Liu, Silver nanoparticles—the real “silver bullet” in clinical medicine, Med.
Chem. Commun., 1 (2010), pp. 125–131
[23] H. Kong, J. Jang, Antibacterial properties of novel poly(methyl methacrylate) nanofiber
containing silver nanoparticles, Langmuir, 24 (2008), pp. 2051–2056
[24] A. Barras, J. Lyskawa, S. Szunerits, P. Woisel, R. Boukherroub, Direct functionalization of
nanodiamond particles using dopamine derivatives, Langmuir, 27 (2011), p. 12451
[25] A. Barras, S. Szunerits, L. Marcon, N. Monfilliette-Dupont, R. Boukherroub, Functionalization of
diamond nanoparticles using “click” chemistry, Langmuir, 26 (2010), pp. 13168–13172
[26] Y.-R. Chang, H.-Y. Lee, K. Chen, C.-C. Chang, D.-S. Tsai, C.-C. Fu, T.-S. Lim, Y.-K. Tzeng, C.-Y. Fang,
C.-C. Han, H.-C. Chang, W. Fann, Mass production and dynamic imaging of fluorescent
nanodiamonds, Nat. Nanotechnol., 3 (2008), pp. 284–288
[27] S.A. Dahoumane, M.N. Nguyen, A. Thorel, J.P. Boudou, M.M. Chehimi, C. Mangeney, Proteinfunctionalized hairy diamond nanoparticles, Langmuir, 25 (2009), pp. 9633–9638
[28] A. Krüger, Hard and soft: biofunctionalized diamond, Angew. Chem. Int. Ed., 45 (2006), pp.
6426–6427
[29] A. Krüger, New carbon materials: biological applications of functionalized nanodiamond
materials, Chem. Eur. J., 14 (2008), pp. 1382–1390
[30] Y. Liang, M. Ozawa, A. Krueger, A general procedure to functionalize agglomerating
nanoparticles demonstrated on nanodiamond, ACS Nano, 3 (2009), pp. 2288–2296
[31] V.N. Mochalin, O.A. Shenderova, D. Ho, Y. Gogotsi, The properties and applications of
nanodiamonds, Nat. Nanotechnol., 7 (2012), p. 11
[32] L. Marcon, F. Riquet, D. Vicogne, S. Szunerits, J.-F. Bodart, R. Boukherroub, Cellular and in vivo
toxicity of functionalized nanodiamond in Xenopus embryos, J. Mater. Chem., 20 (2010), pp. 8064–
8069
[33] K.K. Liu, C.L. Cheng, C.C. Chang, J.I. Chao, Biocompatible and detectable carboxylated
nanodiamond on human cell, Nanotechnology, 18 (2007), p. 325102
[34] A.M. Schrand, H. Huang, C. Carlson, J.J. Schlager, E. Omacr Sawa, S.M. Hussain, L. Dai, Are
diamond nanoparticles cytotoxic?, J. Phys. Chem. B, 111 (2007), pp. 2–7
[35] S.J. Yu, M.W. Kang, H.C. Chang, K.M. Chen, Y.C. Yu, Bright fluorescent nanodiamonds: no
photobleaching and low cytotoxicity, J. Am. Chem. Soc., 127 (2005), pp. 17604–17605
[36] E. Perevedentseva, C.-Y. Cheng, P.-H. Chung, J.-S. Tu, T.-H. Hsieh, C.-L. Cheng, The interaction of
the protein lysozyme with bacteria E. coli observed using nanodiamond labelling, Nanotechnology,
18 (2007), p. 315107
[37] J. Wehling, R. Dringer, R.N. Zare, M. Maas, K. Rezwan, Bactericidal activity of partially oxidized
nanodiamonds, ACS Nano, 8 (2014), p. 6475
[38] M. Hartmann, P. Betz, Y. Sun, S.N. Gorb, T.K. Lindhorst, A. Krueger, Saccharide-modified
nanodiamond conjugates for the efficient detection and removal of pathogenic bacteria, Chem. Eur.
J., 18 (2012), p. 6485
[39] X.R. Wu, T.T. Sun, J.J. Medina, In vitro binding of type 1-fimbriated Escherichia coli to uroplakins
Ia and Ib: relation to urinary tract infections, Proc. Natl. Acad. Sci. U. S. A., 93 (1996), pp. 9630–9635
[40] C.-C. Lin, Y.-C. Yeh, C.-Y. Yang, C.-L. Chen, G.-F. Chen, C.-C. Chen, Y.-C. Wu, Selective binding of
mannose-encapsulated gold nanoparticles to type 1 pili in Escherichia coli, J. Am. Chem. Soc., 124
(2002), p. 3508
[41] T. Patel, Y. Ishiuji, G. Yosipovitch, Menthol: a refreshing look at this ancient compound, J. Am.
Acad. Dermatol., 57 (2007), pp. 873–878
[42] J.A. Farco, O. Grundmann, Menthol — pharmacology of an important naturally medicinal “cool”,
Mini-Rev. Med. Chem., 13 (2013), pp. 124–131
[43] G.P.P. Kamatou, I. Vermaak, A.M. Viljoen, B.M. Lawrence, Menthol: a simple monoterpene with
remarkable biological properties, Phytochemistry, 96 (2013), pp. 15–25
[44] D. Trombetta, F. Castelli, M.G. Sarpietro, V. Venuti, M. Christiani, C. Daniele, A. Saija, G.
Mazzanti, G. Bisignani, Mechanism of antibacterial action of three monoterpenes, Antimicrob.
Agents Chemother., 49 (2005), pp. 2474–2478
[45] G. Iscan, N. Kirimer, M. Kurkcuoglu, K.H. Baser, F. Demirci, Antimicrobial screening of Mentha
piperita essential oils, J. Agric. Food Chem., 50 (2002), pp. 3943–3946
[46] R. Kotan, S. Kordali, A. Cakir, Screening of antibacterial activities of twenty-one oxygenated
monoterpenes, Z. Naturforsch. C, 62 (2007), pp. 507–513
[47] S. Dalleau, E. Cateau, T. Bergès, J.-M. Berheaud, C. Imbert, In vitro activity of terpenes against
Candida biofilms, Int. J. Antimicrob. Agents, 31 (2008), pp. 572–576
[48] A. Tiraferri, C.D. Vecitis, M. Elimelech, Covalent binding of single-walled carbon nanotubes to
polyamide membranes for antimicrobial surface properties, ACS Appl. Mater. Interfaces, 3 (2011),
pp. 2869–2877
[49] S.A. Al-Bataineh, R. Luginbuehl, M. Textor, M. Yan, Covalent immobilization of antibacterial
furanones via photochemical activation of perfluorophenylazide, Langmuir, 25 (2009), pp. 7432–
7437
[50] A. Krüger, Y. Liang, G. Jarre, J. Stegk, Surface functionalisation of detonation diamond suitable
for biological applications, J. Mater. Chem., 16 (2006), p. 2322
[51] V. Pichot, M. Comet, E. Fousson, C. Baras, A. Senger, F. Le Normand, D. Spitzer, An efficient
purification method for detonation nanodiamonds, Diamond Relat. Mater., 17 (2008), p. 13
[52] D.J. Diekema, M.A. Pfaller, F.J. Schmitz, J. Smayevsky, J. Bell, R.N. Jones, M. Beach, Survey of
infections due to Staphylococcus species: frequency of occurrence and antimicrobial susceptibility of
isolates collected in the United States, Canada, Latin America, Europe, and the Western Pacific
Region for the SENTRY Antimicrobial Surveillance Program, 1997–1999, Clin. Infect. Dis., 32 (2001),
pp. S114–S132
[53] E. Boakes, A.M. Kearns, M. Ganner, C. Perry, R.L. Hill, M.J. Ellington, Distinct bacteriophages
encoding panton-valentine leukocidin (PVL) among international methicillin-resistant Staphylococcus
aureus clones harboring PVL, J. Clin. Microbiol., 49 (2010), pp. 684–692
[54] J.P. Zhang, S. Normark, Induction of gene expression in Escherichia coli after pilus-mediated
adherence, Science, 273 (1996), pp. 1234–1236
[55] A. Chwalibog, E. Sawosz, A. Hotowy, J. Szeliga, S. Mitura, K. Mitura, M. Grodzik, P. Orlowski, A.
Sokolowska, Visualization of interaction between inorganic nanoparticles and bacteria or fungi, Int. J.
Nanomedicine, 5 (2010), pp. 1085–1094